Proteases represent a number of families of proteolytic enzymes that catalytically hydrolyze peptide bonds. Principal groups of proteases include metalloproteases, serine proteases, cysteine proteases and aspartic proteases. Proteases, in particular serine proteases, are involved in a number of physiological processes such as blood coagulation, fertilization, inflammation, hormone production, the immune response and fibrinolysis.
Numerous disease states are caused by and can be characterized by alterations in the activity of specific proteases and their inhibitors. For example emphysema, arthritis, thrombosis, cancer metastasis and some forms of hemophilia result from the lack of regulation of serine protease activities (see, for example, Textbook of Biochemistry with Clinical Correlations, John Wiley and Sons, Inc. N.Y. (1993)).
Similarly, proteases have been implicated in cancer metastasis. Increased synthesis of the protease urokinase has been correlated with an increased ability to metastasize in many cancers. Urokinase activates plasmin from plasminogen which is ubiquitously located in the extracellular space and its activation can cause the degradation of the proteins in the extracellular matrix through which the metastasizing tumor cells invade. Plasmin can also activate the collagenases thus promoting the degradation of the collagen in the basement membrane surrounding the capillaries and lymph system thereby allowing tumor cells to invade into the target tissues (Dano, et al. Adv. Cancer. Res., 44: 139 (1985).
Clearly measurement of changes in the activity of specific proteases is clinically significant in the treatment and management of the underlying disease states. Proteases, however, are not easy to assay. Typical approaches include ELISA using antibodies that bind the protease or RIA using various labeled substrates. With their natural substrates assays are difficult to perform and expensive. With currently available synthetic substrates the assays are expensive, insensitive and nonselective. In addition, many "indicator" substrates require high quantifies of protease which results, in part, in the self destruction of the protease.
Recent approaches to protease detection rely on a cleavage-induced spectroscopic change in a departing chromogen or fluorogen located in the P1' position (the amino acid position on the carboxyl side of the cleavable peptide bond) (see, for example U.S. Pat. Nos. 4,557,862 and 4,648,893). However, many proteases require two or three amino acid residues on either side of the scissile bond for recognition of the protease and thus, these approaches lack protease specificity.
Recently however, fluorogenic indicator compositions have been developed in which a "donor" fluorophore is joined to an "acceptor" chromophore by a short bridge containing a (7 amino acid) peptide that is the binding site for an HIV protease and linkers joining the fluorophore and chromophore to the peptide (Wang et al. Tetra. Letts. 45: 6493-6496 (1990)). The signal of the donor fluorophore was quenched by the acceptor chromophore through a process of resonance energy transfer. Cleavage of the peptide resulted in separation of the chromophore and fluorophore, removal of the quench and a subsequent signal from the fluorophore.
Unfortunately, the design of the bridge between the donor and the acceptor led to relatively inefficient quenching limiting the sensitivity of the assay. In addition, the chromophore absorbed light in the ultraviolet range reducing the sensitivity for detection in biological samples which typically contain molecules that absorb strongly in the ultraviolet.
Clearly fluorogenic protease indicators that show a high signal level when cleaved, and a very low signal level when intact, that show a high degree of protease specificity, and that operate exclusively in the visible range thereby rendering them suitable for use in biological samples are desirable. The compositions of the present invention provide these and other benefits.